edited by karl esser the mycota takeshita... · a comprehensive treatise on fungi as experimental...
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Edited byKarl Esser
Biology of the Fungal Cell
A Comprehensive Treatise on Fungi as Experimental Systems for Basic and Applied Research
THE MYCOTA
Dirk Hoffmeister Markus GresslerVolume Editors
VIIIThird Edition
The Cytoskeleton and Polarity Markers During Polarized Growth ofFilamentous Fungi
NORIO TAKESHITA1, REINHARD FISCHER2
CONTENTS
I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 43II. The Actin Cytoskeleton . . . . . . . . . . . . . . . . . . . . . 44III. Spitzenkorper . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 46IV. Septins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 46V. The Microtubule Cytoskeleton . . . . . . . . . . . . . . 47VI. Cell-End Markers for Polarity
Maintenance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49VII. Rho GTPase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51VIII. Cell-End Markers for Polarity
Establishment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51IX. Cell-End Markers for Polarity Focusing . . . . 52X. Transient Polarity Marker Assembly . . . . . . . 54XI. Oscillatory Fungal Cell Growth . . . . . . . . . . . . . 56XII. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 56
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 57
I. Introduction
Maintaining cell polarity is essential for cells toensure homeostasis and their proper function-ing (Goehring and Grill 2013; Wu and Lew2013). Symmetry breaking is often precededby cytoskeleton-dependent polarization of cer-tain key proteins as observed in epithelial cellswith apical-basal polarity, neuronal differentia-tion from dendrites to axons, and migratingcells. Filamentous fungi are highly polarizedeukaryotic cells, which continuously elongatetheir hyphae at the tips. Some distance backfrom the tip, hyphae can initiate new sites of
polar growth in the process of branch forma-tion. The establishment and maintenance ofpolar growth is one fascinating question inbiology. Filamentous fungi are widely used asmodel systems for the analysis of the relation-ship between cell polarity and shape (Harris2006; Fischer et al. 2008; Riquelme et al. 2011,2018; Takeshita et al. 2014). Some filamentousfungi are pathogenic to animals and plants, andoften growth in the host is accompanied by achange from hyphal growth to yeast-likegrowth or vice versa (dimorphism) (Garcia-Vidal et al. 2013). Other fungi are useful inbiotechnology, such as enzyme productionand fermentation in food industry due to theirhigh ability of enzyme secretion (Punt et al.2011). Thus, the analysis of polarized growthof filamentous fungi can contribute to the med-ical, agricultural, and biotechnological fields.
The filamentous ascomycete Aspergillusnidulans has been employed worldwide formore than 60 years as a model organismbecause it is closely related to clinically andeconomically important Aspergilli and it is eas-ily manipulated in the laboratory. The mostcharacteristic cell type of filamentous fungi isthe vegetative hypha. This nonspecialized, syn-cytial (multinucleated) cell is characterized by acontinuous polarized growth mode, mediatedat the tip through the addition of new materialthat is transported from distal regions. Insingle-cell yeasts, such as in budding yeast Sac-charomyces cerevisiae and in fission yeast Schi-zosaccharomyces pombe, polarized growth isrestricted to certain times during the cellcycle, whereas in filamentous fungi cell exten-sion is a continuous and indefinite process.
1Microbiology Research Center for Sustainability (MiCS),Faculty of Life and Environmental Sciences, University ofTsukuba, Tsukuba, Japan; e-mail: [email protected] Department of Microbiology, Institute for Applied Bio-sciences, Karlsruhe Institute of Technology (KIT), Karlsruhe,Germany; e-mail: [email protected]
Biology of the Fungal Cell, 3rd EditionThe Mycota VIIID. Hoffmeister, M. Gressler (Eds.)© Springer Nature Switzerland AG 2019
The extension of hyphal tips requires thecontinuous enlargement of the cell membraneand the extension of the cell wall. Both areachieved through massive vesicle fusion at thetip. The vesicles transport cell wall-synthesizingenzymes and provide new membrane. Vesicletransport as well as all other dynamic processesrelated to polar growth, such as organelle dupli-cation and distribution, or the transport ofRNA, proteins, or lipids, requires cytoskeletalelements. Many of those components were dis-covered early on in mutant screenings followedby genetic and later molecular biological ana-lyses. For instance, tubulin was discoveredthrough a combination of biochemical analyseswith the analysis of mutants with altered sensi-tivity against the microtubule drug benomyl(benA) in A. nidulans (Oakley 2004). Mitoticelements were isolated as temperature-sensitivemutants with a block-in-mitosis (bimA, bimC,bimE) or mutants, which never entered mitosis(never-in-mitosis, nimA, nimX, nimT) (Orr andRosenberger 1976; Morris and Enos 1992;Osmani and Mirabito 2004). In other mutantsof this screening, nuclear distribution (nudA,nudE) was affected (Meyer et al. 1988). Insubsequent suppressor screenings using thebenA33 mutant, tubA and mipA were discov-ered (Morris et al. 1979; Oakley and Oakley1989). The genes encoded beta tubulin (benA),alpha tubulin (tubA), or gamma tubulin(mipA), dynein (nudA), or kinesin (bimC).The mutagenesis approaches performed in S.cerevisiae and S. pombe but also filamentousfungi such as A. nidulans revealed a wealth ofinformation, which could probably not havegenerated by other means. The improvedmolecular biological methods and the genomeinformation opened the possibility of reverse-genetic approaches. With these approaches therole of proteins of conserved pathways wasstudied in other organisms and organism-specific functions were discovered. However,recent major advances in cost-effectivesequencing of entire genomes have the greatpotential to revolutionize our approachesagain and allow intelligent mutant screeningfollowed by bulk sequencing of mutant gen-omes. This strategy reduced mutant analysisfrom months or years to weeks or months(Tan et al. 2014).
Polarized growth is thus studied by genetic,molecular biological, biochemical, and cellbiological methods. However, this research fieldhas benefited more than others from the combi-nation of the still ongoing improvement of themicroscopic techniques and the development offluorescent reporter proteins in recent years.Several overviews have summarized differentaspects of polarized growth in fungi (Changand Peter 2003; Harris and Momany 2004;Penalva 2010; Berepiki et al. 2011; Steinberg2011, 2014; Sudbery 2011; Egan et al. 2012a).
Here we review recent findings unravelingthe mechanism of polarized growth with specialemphasis on the roles of the actin and microtu-bule (MT) cytoskeletons, polarity markers link-ing the two cytoskeletons.
II. The Actin Cytoskeleton
The actin cytoskeleton plays a central role incell morphology of eukaryotic cells (Domin-guez and Holmes 2011). Actin filaments (F-actin), which are composed of linear polymersof G-actin subunits, generate force against theplasma membrane and also act as tracks formyosin motors. The dynamic cycles of poly-merization and depolymerization of G-actinand F-actin are involved in many different keycellular processes, such as cell motility, cytoki-nesis, secretion, and the control of cell mor-phology (Michelot and Drubin 2011).
There are three high-order F-actin struc-tures with distinct functions in filamentousfungi: actin rings, patches, and cables (Berepikiet al. 2011). Studies using anti-actin chemicalagents confirmed that a polymerized actin cyto-skeleton is required for normal apical growthand hyphal tip shape in different fungal organ-isms (Torralba et al. 1998). Phalloidin conju-gated to fluorescent dyes has been widely usedfor imaging F-actin in eukaryotic cells includ-ing fungi such as budding yeast (Amberg 1998),fission yeast (Pelham and Chang 2001), andAshbya gossypii (Walther and Wendland 2004)but does not work in most filamentous fungi(Brent Heath et al. 2003). The immunostainingof actin using an anti-actin antibody and GFP-labeled actin or an actin-binding protein (AbpA
44 N. Takeshita and R. Fischer
in A. nidulans) revealed the localization of theF-actin structures, mainly actin rings and actinpatches, in filamentous fungi (Araujo-Bazanet al. 2008). However, visualization of actincables was difficult by these methods. Recently,specific markers for actin cables, such as Lifeactand tropomyosin, were developed to visualizethem (Taheri-Talesh et al. 2008; Berepiki et al.2010; Delgado-Alvarez et al. 2010). Lifeact,which consists of 17 amino acids from the N-terminus of Abp140p of S. cerevisiae, has beenshown to be a marker for F-actin binding andlabeling in vitro and in yeast cells (Asakuraet al. 1998; Riedl et al. 2008).
The actin rings in cooperation with class IImyosin function in septum formation (Taheri-Talesh et al. 2012; Delgado-Alvarez et al. 2014)(Fig. 1a). Septum formation proceeds accordingto the following series of steps: actin and myo-sin tangle assembly at a septation site, contrac-
tion of the actin ring (or called actomyosinring), actin-mediated invagination of theplasma membrane, and deposition of the chi-tinous primary septum (Momany and Hamer1997; Delgado-Alvarez et al. 2014). Mutantanalysis of class II myosin (myoB mutation inthe converter subdomain in A. nidulans) sug-gests that the motor activity is necessary for thecontraction of the actin ring (Hill et al. 2015).The actin ring assembly and septum formationare controlled through the nuclear position andcell cycle progression in A. nidulans (Harris2001).
Actin patches are peripheral punctate struc-tures where probably the endocytic machinerylocalizes (Araujo-Bazan et al. 2008) (Fig. 1b). Thepredominant localization of these patches at sub-apical regions suggests spatial coupling of apicalexocytosis and subapical compensatory endocy-tosis (Penalva 2010). Class I myosins function in
Fig. 1 Distinct roles of actin cytoskeletons and relatedmyosin in A. nidulans. (a) Actin ring and myosin II(actomyosin ring) for septation. (b) Actin patch andmyosin I for endocytosis. (c) Actin cables and myosin
V for exocytosis. Scheme of tip growth in A. nidulanshyphae. Vesicle trafficking via the actin and MT cyto-skeleton. Before fusion with the plasma membrane,secretion vesicles accumulate at Spitzenkorper
The Cytoskeleton and Polarity Markers During Polarized Growth of Filamentous Fungi 45
endocytosis to support invagination of endocyticvesicles (Kaksonen et al. 2006). Mutant analysisof myoA, the class I myosin in A. nidulans,revealed that the function of MyoA in endocyto-sis is regulated through phosphorylation by amember of the p21-activated kinase (PAK) fam-ily (Yamashita and May 1998). The mutant phe-notype of genes involved in endocytosis indicatesthat endocytosis is essential (Araujo-Bazan et al.2008). Besides the internalization of extracellularmolecules, plasma membrane proteins, andlipids by endocytosis, endocytic recycling ofpolarizedmaterial at the hyphal tip and a balancebetween endocytosis and exocytosis at thehyphal tip are assumed to control polarizedgrowth and cell shape (Shaw et al. 2011).
Actin cables are linear bundles of shortactin filaments nucleated by formins that arepresent at the apex of the hyphae. As mentionedbefore, dynamic actin cables are generally verydifficult to visualize; however, recently specificmarkers, such as Lifeact and tropomyosin, weredeveloped (Taheri-Talesh et al. 2008; Berepikiet al. 2010; Delgado-Alvarez et al. 2010). In N.crassa, Lifeact has been used to visualizedynamic actin cables and patches (Berepikiet al. 2010; Delgado-Alvarez et al. 2010). How-ever, it has to be considered that overexpres-sion of the construct may cause some artifacts(Bergs et al. 2016). Tropomyosin is a conservedactin filament-binding protein and regulatesthe interaction between actin and myosin inresponse to Ca2+ (Gunning et al. 2005). Tropo-myosin has been used as a marker for actincables in A. nidulans and N. crassa (Evangelistaet al. 2002; Pearson et al. 2004; Taheri-Taleshet al. 2008). GFP-labeled tropomyosin, TpmA inA. nidulans, revealed the dynamic behaviorwith cycles of elongation and shrinkage(Fig. 1c) (Bergs et al. 2016). Multiple actincables were formed from the hyphal tip witheach actin cable showing elongation andshrinkage in an independent manner.
Actin cables are present at the apex of hyphae and arethought to serve as tracks for class V myosin-dependent secretory vesicle transport to the tip(Fig. 1c) (Taheri-Talesh et al. 2008, 2012; Berepikiet al. 2011; Pantazopoulou et al. 2014). The “basic”growth machinery involved in the formation of actin
cables, vesicle transport, and exocytosis, such as for-min, the polarisome (protein complex Spa2, Pea2,Aip3/Bud6, and formin Bni1 in S. cerevisiae), class Vmyosin, and the exocyst complex (octameric proteincomplex involved in the tethering of post-Golgi vesiclesto the plasma membrane prior to vesicle fusion), isrelatively conserved among eukaryotic cells and loca-lized to the apex of hyphae (Harris et al. 2005; Sudbery2011). Maturation of late Golgi cisternae into exocyticpost-Golgi carriers was visualized in A. nidulans (Pan-tazopoulou et al. 2014). These carriers move on a MT-based bidirectional conveyor belt relaying them toactin, which ultimately focuses exocytosis at the apex.
III. Spitzenkorper
Before fusion, the secretion vesicles accumulateat the hyphal tip in a structure called Spitzen-korperor vesicle supply center (VSC) (Grove andBracker 1970; Harris et al. 2005), a special struc-ture in filamentous fungi, which determinesgrowth direction of the hyphae (Bartnicki-Garciaet al. 1995; Riquelme and Sanchez-Leon 2014;Riquelme et al. 1998, 2014) (Fig. 1c). A VSC inmotion provides a rational basis to predict howthe secretory apparatus generates morpho-genesis. The Spitzenkorper is believed to func-tion as a VSC that regulates the delivery of cellwall-building vesicles to the apical cell surface,since the simulation analysis of VSC advance as aSpitzenkorper enables to mimic the hyphalgrowth of N. crassa (Bartnicki-Garcıa et al.1989; Riquelme et al. 2000). The exact compo-sition and organization of the Spitzenkorper hasbeen vigorously elucidated (Riquelme et al. 2007,2014; Sanchez-Leon et al. 2011; Fajardo-Someraet al. 2015). In N. crassa, all chitin synthaseslocalize at the Spitzenkorper core, whereasmacrovesicles carrying a b-1,3-glucan synthasecomplex occupy the Spitzenkorper outer layer.Similar spatial distribution of DnfA and DnfB,two P4 ATPases, in the Spitzenkorper wasobserved inA. nidulans (Schultzhaus et al. 2015).
IV. Septins
As filament-forming proteins, septins can beconsidered as a part of the cytoskeleton. Septinsare a conserved family of GTP-binding proteins
46 N. Takeshita and R. Fischer
that form filaments in fungi and animals. Dif-ferent septins form protein complexes witheach other and form heteropolymers showinga variety of higher-order structures (Mostowyand Cossart 2012). In S. cerevisiae, five septinproteins (Cdc3, Cdc10, Cdc11, Cdc12, Shs1)form an hourglass structure associated withthe plasma membrane at the mother-bud neck(Bertin et al. 2012). They act as a scaffold forproteins involved in cell division (Oh and Bi2011). In filamentous fungi, in addition to sep-tum formation, septins have been shown tohave roles in morphogenesis, coordinatingnuclear division, and organizing the cytoskele-ton (Lindsey and Momany 2006; Gladfelter2010; Bridges and Gladfelter 2014).
In the hyphal form of the human pathogen Candidaalbicans, septins form three distinct structures, a dif-fuse band at the base of hyphae, a bright double ring atseptation sites, and a diffuse cap at hyphal tips (Sudb-ery 2001; Warenda and Konopka 2002). In the filamen-tous phytopathogen A. gossypii, septins form a diffusecap at hyphal tips and rings composed of discrete barsat septation sites and at newly emerging branches (Hel-fer and Gladfelter 2006; DeMay et al. 2009). In C. albi-cans and A. gossypii, the diffuse cap initially appears atthe hyphal tip and travels with the hyphal tip until anunknown signal triggers detachment from the tip andthe formation of an anchored, higher-order ring. Thehigher-order ring encircles the hyphal cell cortex andpersists at that location, while the tip then continues togrow. Septation sites are determined by positions of thehigh-order ring that detaches from the tip and anchorsat the hyphal cell cortex in these fungi. The structuralchange of septins is regulated through their phosphor-ylation by cyclin-dependent kinases in C. albicans(Sinha et al. 2007) or septin-associated kinases in A.gossypii (DeMay et al. 2009).
In A. nidulans and N. crassa, septins form differenthigher-order structures, rings, filaments, bars, bands,and caps (Westfall and Momany 2002; Lindsey et al.2010; Berepiki and Read 2013; Hernandez-Rodriguezet al. 2014). They are important for septation, germina-tion, branch emergence, and asexual spore formation.In the basidiomycete plant pathogen Ustilago maydis,septins localize to a variety of structures, collars at thebud neck and filaments at growing cell tips that runalong the length of the cell and partially colocalize withMTs (Boyce et al. 2005; Alvarez-Tabares and Perez-Martin 2010). Septin filaments in the basidiomycetedikaryotic hyphae of Cryptococcus neoformans havealso been shown to occasionally colocalize with MTs(Kozubowski and Heitman 2010). These septins haveroles in morphogenesis and host infection. In the plantpathogen Magnaporthe oryzae, the location of the
appressorium septum is determined by the site of sep-tin ring assembly (Saunders et al. 2010). The septin ringfunctions as a scaffold for proteins required for appres-sorium formation (Dagdas et al. 2012). The recentreports on septins in filamentous fungi have revealednew roles for these cytoskeletal polymers.
V. The Microtubule Cytoskeleton
MTs play a crucial role during mitosis but servealso additional functions in interphase in fila-mentous fungi. They are important for the dis-tribution of nuclei and other organelles andserve as tracks for endosomes and other vesi-cles; thus they are important for rapid hyphalgrowth (Xiang and Fischer 2004; Horio andOakley 2005; Egan et al. 2012a; Steinberg 2014).
The rather stable minus end of MTs islocated at the MT-organizing center (MTOC),whereas the plus end is facing to the cell periph-ery with alternate growing and shrinking phases.In filamentous fungi, spindle pole bodies (SPBs)serve as MTOCs (Fig. 2a) (Oakley et al. 1990).They contain g-tubulin, first discovered in A.nidulans, which is required for nucleation ofMTs (Oakley and Oakley 1989; Oakley et al.1990). Furthermore, there is good evidence thatareas close to the septa act as MTOCs in A.nidulans (sMTOCs) (Fig. 2a) (Veith et al. 2005;Xiong and Oakley 2009; Zekert and Fischer 2009;Zhang et al. 2017). The composition of thoseMTOCs, their role, and their tethering to theseptal membrane remain to be elusive. However,there is also good evidence for several MTOCs inS. pombe, in U. maydis, and in higher eukar-yotes. In S. pombe equatorial (eMTOC) andinterphase MTOCs (iMTOC) have beendescribed (Sawin and Tran 2006) but appear tobe only temporally active during certain stagesof the cell cycle, and their exact nature also isenigmatic. In higher eukaryotes the Golgi appa-ratus had some MT-forming activity (Sutterlinand Colanzi 2010).
In the tip compartment of A. nidulans, mostMTs are oriented with their dynamic plus endstoward the hyphal tip (Fig. 2a, b) (Konzack et al.2005). There are only a few MTs found in inter-phase compartments, and nuclei migrate proba-bly along MTs until they reach a certain
The Cytoskeleton and Polarity Markers During Polarized Growth of Filamentous Fungi 47
position. The entire hypha looks therefore veryorganized with evenly spaced nuclei.
Two classes of MT-dependent motors, theminus end-directed dynein and the plus end-directed kinesins, are involved in positioning oforganelles and transport of membranes.Whereas genomes of filamentous fungi containa single dynein motor, they usually encode 10–12 kinesins (Schoch et al. 2003). The function ofkinesin-3 and the dynein motor in the transportof early endosomes have been analyzed deeply(Fig. 2c) (Steinberg 2011, 2014; Egan et al.2012a). In A. nidulans hyphae, most MTsbetween the hyphal tip and the most proximalnucleus were polarized with their plus endsoriented toward the growing hyphal tip. Dyneinand kinesin-3/UncA are the opposite motorsresponsible for bidirectional transport of endo-somes and peroxisomes (Abenza et al. 2009;
Egan et al. 2012b). Studies of A. nidulanskinesin-3 implicated indirect evidence for theexistence of a subpopulation of detyrosinatedMTs (Zekert and Fischer 2009; Seidel et al.2012). However, a final proof for the existenceof posttranslationally modified tubulin in fungiis yet still missing.
Endocytic recycling at subapical regionssupports fungal tip growth. The bidirectionalmotility of early endosomes is thought to beinvolved in sorting the endocytic cargo to thesubapical vacuole for degradation (Wedlich-Soldner et al. 2000). An unexpected new rolefor early endosome motility was revealed in U.maydis. The RNA-binding protein Rrm4 trans-ports various mRNAs on moving early endo-somes, suggesting that early endosome motilitytoward the cell ends supports polar delivery ofRrm4-bound RNAs (Konig et al. 2009; Bau-
Fig. 2 Microtubule cytoskeleton in A. nidulans. (a)MT-organizing center (MTOC) at spindle pole bodiesand septa. Most MTs are oriented with their dynamicplus ends toward the hyphal tip in the tip compartment.
(b) Time lapse of GFP-KipA (MT plus end marker) andmCherry-TubA (MT) in A. nidulans hyphae. (c) Imageof GFP-RabA (early endosome marker) and mCherry-TubA (MT) and kymograph of GFP-RabA
48 N. Takeshita and R. Fischer
mann et al. 2012). In addition, it was shown thatthe mRNA of the septins cdc3 and cdc12 isactively translated on moving early endosomes,and both proteins bind to early endosomes(Baumann et al. 2014). In fact, it was confirmedthat ribosomes attach to early endosomes andare translationally active during the transport(Higuchi et al. 2014). It is suggested that bidi-rectional early endosome motility constantlystirs the translation machinery, which diffusespassively in the cytoplasm and thereby contri-butes to distribute the ribosomes throughoutthe cell (Steinberg 2014; Haag et al. 2015).Indeed, endosomal transport of heteromericseptin complexes along microtubules is crucialfor formation of higher-order structures in U.maydis (Zander et al. 2016).
The deletion of conventional kinesin (kinesin-1)decreased the growth rate and caused defects in Spit-zenkorper stability, protein secretion, and pathogenic-ity (Lehmler et al. 1997; Seiler et al. 1997, 1999; Requenaet al. 2001; Schuster et al. 2012). These results suggest apossible conserved role in vesicle transportation simi-lar to higher eukaryotic cells. High-speed imagingrevealed vesicle transport of chitin synthases (Take-shita et al. 2015). The frequency of the transport wasclearly decreased in the absence of kinA, kinesin-1 in A.nidulans. Secretory vesicles are thought to be trans-ported by kinesin-1 along MTs for long distancestoward hyphal tips in filamentous fungi, although thelocalization of the ER and the Golgi close to hyphal tipsraises questions about the function and cargo ofkinesin-1 (Markina-Inarrairaegui et al. 2013; Pinaret al. 2013). Possibly long-distance transport of secre-tion vesicles is less important and that actin-dependentmovement is rather sufficient. Indeed, hyphal extensioncan occur quite long without functional MTs but isimmediately stopped as soon as the integrity of theactin cytoskeleton is disturbed (Torralba et al. 1998;Horio and Oakley 2005). Although the role of MTsand the different cytoskeletons could be diverse indifferent fungi, vesicle movement and delivery to thetip plasma membrane likely depend on the cooperationof actin- and MT-dependent motors (Zhang et al. 2011;Schuster et al. 2012; Taheri-Talesh et al. 2012).
VI. Cell-End Markers for PolarityMaintenance
Cell polarity is essential for the proper func-tioning of many cell types. During cellular mor-
phogenesis—from fission yeast to humancells—MTs deliver positional information tothe proper site of cortical polarity (Siegristand Doe 2007; Li and Gundersen 2008). Thepolarization of the actin cytoskeleton and sig-nal transduction cascades and continuousmembrane transport toward the growth sitedepend on MTs.
Because MT dynamics and many MT func-tions are conserved among eukaryotes, lowereukaryotes can serve as excellent models. In S.pombe, the kelch-repeat protein Tea1 is deliv-ered by growing MTs to the cell ends (Fig. 3a)(Mata and Nurse 1997). Tea1 reaches the MTplus end with the kinesin-7, Tea2 (Browninget al. 2000, 2003), and is anchored at the cellend membrane through the interaction with theprenylated protein, Mod5 (Snaith and Sawin2003). At the cell end, Tea1 interacts with addi-tional components, which ultimately recruit theformin For3 (Martin and Chang 2003; Feier-bach et al. 2004; Tatebe et al. 2008). For3forms actin cables required for exocytosis andpolarized growth. Cell-end markers, Tea1 andMod5, thus transmit positional informationregulated by MTs to the actin cytoskeletonand thereby contribute to polarized growth.The cell-end marker genes were identifiedafter analysis of polarity mutants (T-shaped orbent cells). The mutants of tea1 and tea2 (tipelongation aberrant) and mod5 (morphologydefective) exhibit T-shaped or bent cells as aresult of the mislocalization of the polarity siteaway from the center of the cell end.
The molecular mechanism of MTs to regu-late polarity maintenance is principally con-served from S. pombe to filamentous fungi(Fig. 3a) (Riquelme et al. 1998; Fischer et al.2008; Takeshita et al. 2008; Higashitsuji et al.2009; Takeshita and Fischer 2011). Tea1 andTea2 are conserved in A. nidulans as TeaA(Tea1) and KipA (Tea2), respectively (Konzacket al. 2005; Takeshita et al. 2008). Although theMod5 sequence is not conserved in filamentousfungi, a functional counterpart, TeaR, was dis-covered by screening for proteins that harbor aC-terminal prenylation motif in A. nidulans(Takeshita et al. 2008). TeaA is conserved inAscomycetes and some Basidiomycetes, such asCryptococcus neoformans and Puccinia graminis,
The Cytoskeleton and Polarity Markers During Polarized Growth of Filamentous Fungi 49
but not in Zygomycetes. TeaR is generally con-served in Ascomycetes except in Hemiascomy-cetes. The A. nidulans kipA, teaA, and teaRmutants show defects in polarity maintenance,which leads to curved (DkipA orDteaR) or zigzaggrowing hyphae (DteaA) (Fig. 3b). The two cell-end markers, TeaA and TeaR, localize at hyphaltips interdependently (Fig. 3c). TeaA is deliveredto hyphal tips by growing MTs (Takeshita andFischer 2011) and anchored to the hyphal tipcortex though the interaction with TeaR (Take-shita et al. 2008). TeaA, along with TeaC, recruitsthe formin, SepA, to the growth zone (Higashit-suji et al. 2009). The conserved mechanism of“cell-end markers,” TeaA and TeaR, to transmitpositional information from MTs to the actin
cytoskeleton is required for maintenance ofpolarity and the growth direction of hyphae(Fig. 3d). Although the cell-end marker proteinsappear to be conserved in other filamentousfungi, it remains to be studied if the rolesassigned to them in A. nidulans are also con-served. For instance, the MT cytoskeleton ismuch more complex in N. crassa.
MTs in A. nidulans are necessary to maintain the polar-ity at the tip of hyphae through cell-end markers.Besides this function, MTs have additional functionsin filamentous fungi, such as nuclear distribution andthe movement of vesicles and other organelles; thusthey are important for rapid hyphal growth. In contrast,MTs in S. pombe are required for polarity maintenancethrough cell-end markers but are not necessary for
Fig. 3 Role of cell-end markers in A. nidulans. (a)Scheme of the function of cell-end markers in S. pombeand A. nidulans. (b) Differential interference contrastimages of wild type, DteaA, DteaR strains. DteaR strainsexhibited curved hyphae and DteaA strains exhibitedzigzag hyphae. (c) Localization of mRFP1-TeaA and
GFP-TeaR (upper) at the hyphal tips. Bimolecular fluo-rescence complementation (BiFC) assay of TeaA andTeaR at the hyphal tip (lower left). Localization ofmRFP1-TeaA at the hyphal tip and MT plus end (lowerright). (d) Scheme of growth direction regulated by cell-end markers and cell cytoskeletons
50 N. Takeshita and R. Fischer
vesicle trafficking. Actin cables grow toward the grow-ing cell ends, and Myo52, a myosin V, is responsible forpolarized secretion of vesicles along actin cables andhence membrane enlargement and secretion of cellwall-synthesizing enzymes (Montegi et al. 2001; Winet al. 2001; Mulvihill et al. 2006).
VII. Rho GTPase
The small Rho-type GTPases Cdc42 and Rac1are key regulators of eukaryotic cell polarity(Jaffe and Hall 2005; Virag et al. 2007). Theswitching between inactive GDP-bound andactive GTP-bound states is controlled by gua-nine nucleotide exchange factors (GEFs)(Schmidt and Hall 2002) and GTPase-activatingproteins (GAPs) (Bernards and Settleman2004). Active GTPases stimulate multiple effec-tor molecules, such as p21-activated kinases(PAKs), mitogen-activated protein kinases(MAPKs), formin, and subunits of the exocystcomplex, which regulate numerous cellularprocesses including the rearrangement of theactin cytoskeleton, targeted vesicle transport,and exocytosis (Bishop and Hall 2000). In S.cerevisiae, two positive feedback loops arethought to contribute to Cdc42 polarization.One pathway involves recruitment of GEFsand effector complexes from the cytoplasm ina cytoskeleton-independent manner (Johnsonet al. 2011). The other one is a vesicle-recyclingfeedback loop, where Cdc42 orients actincables, which in turn deliver Cdc42 as cargoon secretory vesicles (Wedlich-Soldner et al.2003).
In S. cerevisiae and S. pombe, Rac1 orthologs are notconserved, and Cdc42 alone is necessary and sufficientto control polarized growth (Adams et al. 1990; Millerand Johnson 1994). In contrast, filamentous fungirequire both Rho GTPases to regulate their hyphalgrowth (Mahlert et al. 2006; Virag et al. 2007; Lichiuset al. 2014). Both Cdc42 and Rac1 share at least oneoverlapping function that is required for polarity estab-lishment. The combination of Dcdc42 with Drac1appeared synthetically lethal in A. nidulans (Viraget al. 2007). The cell-end marker deletion strains in A.nidulans lose the axis of polarity, although the hyphaestill continue polarized tip growth. Although the polar-ization of Cdc42 and Rac1 by positive feedback loops asknown in S. cerevisiae remains elusive in filamentous
fungi, the Rho GTPases are possibly involved in polar-ity establishment independently of the cell-end markersin A. nidulans.
VIII. Cell-End Markers for PolarityEstablishment
MTs and cell-end markers are not essential forspore germination itself but only for site selec-tion of germination (Takeshita and Fischer2011). During conidia germination, TeaAappeared at growth sites prior to the appear-ance of a small germination bud, and then MTscontact the cortex of the tip, while the polarizedMTs likely deliver more TeaA and other pro-teins to the bud site, enforcing polar growth.Wild-type spores always germinate at only onesite to form one hypha; in contrast, spores ofDteaA often germinated at two (40%) or threesites (3%) simultaneously (Fig. 4a). A similarphenotype of multi-germtube formation wasobserved using the MT-destabilizing drug ben-omyl. It had been shown already that MTs arenot essential for the germination process itself(Oakley and Morris 1980). TeaA and MTs arenot necessary for the emergence of the germ-tube but probably rather for restricting germi-nation to a specific place.
When the spores start germination, theygrow isotropically at first, and then they switchto polarized growth with new material added tothe tip of an emerging germtube. In A. nidu-lans, temperature-sensitive swo (swollen cell)mutants representing genes involved in polar-ity establishment and polarity maintenancewere isolated (Momany et al. 1999). swoC(putative pseudouridylate synthase) and swoF(N-myristoyl transferase) (Shaw et al. 2002) arerequired to establish polarity, while swoA (O-mannosyltransferase) is required to maintainpolarity. Besides that, the terminal phenotypeof Dcdc42Drac1 indicates their functions arerequired to establish polarity (Virag et al.2007). In addition, the morphological abnorm-alities of mutants involved in endocytosis indi-cate the importance of endocytosis in polarityestablishment and polarity maintenance (Leeet al. 2008; Hervas-Aguilar and Penalva 2010;Shaw et al. 2011).
The Cytoskeleton and Polarity Markers During Polarized Growth of Filamentous Fungi 51
Once the first hypha reaches a determinatelength, a second germtube appears on thespore. This second germination site normallylies opposite of the first hypha (Harris et al.1999). The second germtube appeared afterthe first septum at the base of the first hyphawas formed. TeaA appeared at the second ger-mination site, opposite the first hypha, afterseptation in the first hypha. In A. nidulans,MTs are formed from SPBs and from septalMTOCs (Konzack et al. 2005; Veith et al. 2005;Xiong and Oakley 2009). MTs emanating fromthe septum of the first hypha grew toward thefirst germtube as well as in the direction of thespore and reached the second germination tipwhere TeaA localized (Fig. 4b). MTs originatingfrom the septal MTOCs are thus important forTeaA delivery and may explain the bipolar ger-mination pattern (Takeshita and Fischer 2011).
There appears to be two patterns of lateralbranching: branches associated with septa andrandom branching. In several fungi includingmembers of the Saccharomycotina (A. gossypii),zygomycetes (Basidiobolus ranarum), and basi-diomycetes (Coprinus species), new branchesemerge adjacent to septa (Trinci 1974). A ran-dom pattern of branching is observed in A.
nidulans. Although the cell-end markers local-ize to branching sites prior to branchingemerges, the mechanism of branch site selec-tion remains largely unknown (Fig. 4c) (Harris2008).
IX. Cell-End Markers for PolarityFocusing
The role of MTs in transmitting positionalinformation through delivery of cell-end mar-kers to the growth machinery is conserved inboth S. pombe and A. nidulans. A Spitzenkor-per, however, can only be observed in filamen-tous fungi but not at cell ends of fission yeast(Fig. 5a). This difference could be due to differ-ent growth speeds (Kohli et al. 2008). Anotherpossible reason is that the cell-end markersconcentrate at the apex of hyphae in A. nidu-lans, whereas the cell-end markers localize atmultiple sites along cell ends in fission yeast(Dodgson et al. 2013). The positive feedbackloop defined through the interdependence ofTeaA and TeaR could be important for theirconcentration, but not sufficient because this
Fig. 4 Cell-end markers forpolarity establishment. (a) Ger-mination in wild type, and sev-eral mutants. (b) Scheme ofsecond germination site selec-tion by MTs from septal MTOCand TeaA. (c) Branching siteselection
52 N. Takeshita and R. Fischer
mechanism is conserved in S. pombe as well(Snaith and Sawin 2003; Takeshita et al. 2008;Bicho et al. 2010). MTs in A. nidulans elongatetoward the tips and tend to converge in theapical region (Konzack et al. 2005), which isnot observed in S. pombe. The central positionof TeaA at the tip correlated with the conver-gence of the MT plus ends to a single point. Inthe absence of TeaA, MTs often contacted themembrane off the center of the apex (Fig. 5b)(Takeshita et al. 2008, 2013).
A recent study showed that a functionalconnection between TeaA and theMT polymer-ase AlpA is required for proper regulation ofMT growth at hyphal tips (Takeshita et al.2013). AlpA is a member of the XMAP215/Dis1 family whose conserved TOG domains,which contain multiple HEAT repeats, are
known to bind tubulin from yeast to human(Al-Bassam et al. 2007). XMAP215 from Xeno-pus laevis catalyzes the addition of tubulindimers to the growing plus ends (Brouhardet al. 2008; Al-Bassam and Chang 2011). A.nidulans AlpA decorates MT filaments andaccumulates at MT plus ends (Enke et al.2007). Deletion of alpA resulted in a drasticreduction of the MT array and dynamics. MTin vitro polymerization assays with purifiedtubulin from porcine brains and recombinantAlpA have revealed the activity of AlpA as a MTpolymerase (Fig. 5c) (Takeshita et al. 2013). TheMT growth speed in vitro was comparable withthat of XMAP215 of X. laevis and approxi-mately fourfold higher than that of Alp14, theorthologue in S. pombe (Brouhard et al. 2008;Al-Bassam et al. 2012). The rate of MT
Fig. 5 Cell-end markers for polarity focusing. (a) Com-parison of the localization of cell-end markers and thegrowth machinery in S. pombe and A. nidulans. (b)Behavior of MTs at hyphal tips in A. nidulans wild typeand DteaA strains. (c) In vitro MT polymerization assay.
Images of a seed MT (red) with a dynamic MT latticegrowing from the plus end (green). Kymographs of MTsin the absence of AlpA (right) and presence of 100 nMAlpA (lower). (d) Scheme of the interaction betweenTeaA at the hyphal tip cortex and AlpA at MT plus ends
The Cytoskeleton and Polarity Markers During Polarized Growth of Filamentous Fungi 53
polymerization in vivo in A. nidulans leadinghyphae is approximately threefold higher thanin S. pombe, consistent with the ratio in vitro(Drummond and Cross 2000; Efimov et al.2006). However, AlpA-dependent MT growthspeed in vitro was approximately only half ofthe one determined in vivo (6 mm/min com-pared to 13 ! 3 mm/min). Therefore, othermicrotubule plus-end-tracking proteins arelikely to enhance the AlpA activity for MTgrowth in vivo.
As a difference to S. pombe, A. nidulansTeaA is involved in the convergence of MTplus ends at the tip apex, suggesting specificinteractions of the MT plus end with the cortex.One possibility is an interaction between TeaAand AlpA (Takeshita et al. 2013). MT polymeri-zation assays in vitro showed that TeaAincreased the catastrophe frequency of MTs inthe presence of AlpA, and TeaA reduced thein vitro AlpA activity significantly. From theseresults it was concluded that AlpA promotesMT growth at MT plus ends until MTs reachthe hyphal tip, where TeaA blocks the AlpAactivity and induces MT catastrophe (Fig. 5d).The interdependence of TeaA and MTs couldact as a positive feedback loop to concentrateTeaA at the apex resulting in well-focused vesi-cle secretion for the organization of the Spit-zenkorper (Bartnicki-Garcia et al. 1995;Riquelme et al. 2014).
X. Transient Polarity MarkerAssembly
The membrane-associated cell-end markerTeaR is highly dynamic at growing hyphal tips(Ishitsuka et al. 2015). A MT grows toward thehyphal tip, pauses in close contact with theapical membrane, and then undergoes a catas-trophe event resulting in retraction. TeaR accu-mulates at the hyphal tip membrane and thendecreases immediately after a MT plus-endtouches the tip membrane and starts to shrink.Colocalization studies indicate that TeaR clus-ters represent zones of exocytosis at the apicalmembrane. In general, membrane-bindingpolarity markers are delivered to the plasma
membrane by vesicle transport and exocytosis.After exocytosis, vesicles fuse with the plasmamembrane, leading to its extension. Simulationanalysis had predicted that membrane inser-tion by active exocytosis would dilute and/ordisperse membrane-binding polarity markersduring fast hyphal growth (Savage et al. 2012).There was no clear answer to the question ofhow cell polarity is maintained during inces-sant vesicle exocytosis, especially for rapidlygrowing systems such as filamentous fungi.One of the problems to capture the complexprocess is that conventional live cell imagingmethods lack the resolution.
The localization of TeaR cluster at hyphaltips has been analyzed by super-resolutionmicroscopy PALM (photoactivation localiza-tion microscopy) (Ishitsuka et al. 2015). Theresolution of conventional light microscopytechniques is limited to about 250 nm due tolight diffraction. Super-resolution microscopytechniques, such as PALM, have overcome thediffraction limit, resulting in lateral image res-olution as high as 20 nm and providing a pow-erful tool for the investigation of proteinlocalization in high detail (Sahl and Moerner2013). The PALM imaging analysis revealedthat TeaR transiently assembles (approximately120 nm, 20 TeaR proteins per cluster on aver-age) at the hyphal tip membrane and dispersesalong the membrane after exocytosis, whichinserts a new membrane that results in localmembrane extension (Fig. 6a) (Ishitsuka et al.2015).
These findings gave rise to a “transientpolarity assembly model” to explain how fungaltip cells extend through repeated cycles of TeaRassembly/disassembly, actin polymerization,and exocytosis rather than by constant elonga-tion (Fig. 6b) (Takeshita 2016). The findings ofcolocalization studies support the notion thatTeaR clusters represent zones of exocytosis andare prerequisite for apical membrane exten-sion. In this model, the interaction betweenthe MT plus end, where TeaA is located, andTeaR at the apical membrane initiates therecruitment of other polarity markers, resultingin the assembly of TeaR polarity site. The accu-mulated cell-end markers induce actin poly-merization followed by active exocytosis.
54 N. Takeshita and R. Fischer
Fig.6Oscillatoryfungalcellgrow
th.(a)Tim
e-lapsePALM
ofTeaRshow
sthe
appearance
ofanew
cluster(t¼
5s),a
translationalmovem
ent(t¼
10s),anda
spreadingof
thesignalalon
gtheplasmamem
brane.Lo
calmem
braneextension
(t¼
0in
red,
t¼
22.5
sin
green).(b)Schemeof
transientpo
larity
mod
eland
tempo
rally
controlledactinpo
lymerizationandexocytosiscoordinated
bypu
lsed
Ca2
+influx.(c)Tim
e-lapseclusteranalysisof
PALM
images
ofchitin
synthase
ChsB.(Left)
Localizationim
ageof
ahyph
awithmEosFP
thermo-ChsBclusters
(500
fram
es).
(Right)Sequ
ence
ofChsB
clusterim
ages
(clustersin
different
colors)rend
ered
from
images
reconstructed
bymovingwindo
wbinn
ing(cluster
images
of2.5stimeintervalsforatotalp
eriodof
125s)
The Cytoskeleton and Polarity Markers During Polarized Growth of Filamentous Fungi 55
Newly synthesized TeaR is delivered to the tipmembrane on secretory vesicles through MTand actin cables. The plasma membrane extendslocally at the site of vesicle fusion, and, subse-quently, the TeaR polarity site is dispersed ordisplaced along the membrane. Once the polar-ity site is disassembled, however, nextMT comesto the tip and gathers TeaR floating in the mem-brane through the interaction with TeaA at theMT plus end and the cycle starts over.
In line with this model, recent work on N. crassa hasidentified bursts of exocytotic events at various siteswithin the apical membrane rather than a persistentexocytosis site (Riquelme et al. 2014). Mathematicalsimulation analysis confirmed the validity of the tran-sient polarity assembly model (Ishitsuka et al. 2015).The simultaneous visualization of actin cables and MTssuggests temporally and spatially coordinated polymer-ization and depolymerization between the two cytoske-letons (Bergs et al. 2016). Interaction between MigA, aMT plus-end localizing protein, and a class V myosinsuggests that an active mechanism captures MTs andpulls the ends along actin filaments (Manck et al. 2015).These results also support the model.
XI. Oscillatory Fungal Cell Growth
Many dynamic cellular processes that appearcontinuous are driven by underlying mechan-isms that oscillate with distinct periods. Forexample, eukaryotic cells do not grow continu-ously but rather by pulsed extension of theperiphery. Stepwise cell extension at the hyphaltips of several filamentous fungi was discovered20 years ago (Lopez-Franco et al. 1994), butonly a few molecular details of the mechanismhave been clarified since then. The “transientpolarity assembly model” also implies the step-wise cell extension. Indeed, the time-lapsePALM imaging revealed that the cell extensionrate of hyphae is not constant but varies in anoscillatory manner (Fig. 6c) (Zhou et al. 2018).
In addition, a recent study has providedevidence for molecular mechanism of oscilla-tory fungal cell growth. Live cell imaginganalyses revealed oscillations of actin assemblyand exocytosis in growing hyphal tips (Take-shita et al. 2017). Intracellular Ca2+ levels are
known to regulate actin assembly and vesiclefusion (Janmey 1994; Schneggenburger andNeher 2005). Ca2+ levels also pulsed at thehyphal tips (Kim et al. 2012). The fluores-cence-based Ca2+ biosensor R-GECO variesthe emission according to Ca2+ concentrationsand has enabled the visualization of pulsatileCa2+ concentrations in growing hyphal tips ofA. nidulans (Takeshita et al. 2017). IntracellularCa2+ levels pulse at ~30-second intervals. Thesepositively and temporally correlate withamounts of F-actin and secretory vesicles athyphal tips. Orthologues of Ca2+ channels atthe plasma membrane in A. nidulans arerequired for proper tip growth and the oscilla-tion of F-actin, secretory vesicles, and Ca2+
level pulses (Wang et al. 2012; Takeshita et al.2017). These results suggest that pulsed Ca2+
influx coordinates the temporally controlledactin polymerization and exocytosis that drivestepwise cell extension (Fig. 6b). The oscillatoryfungal growth could be important for dynamicresponses to external and internal signals inchemotropism, cell-cell fusion, microbial inter-action, and the fungal penetration of plant andanimal cells (Takeshita 2018).
XII. Conclusion
The establishment and maintenance of cellpolarity in fungi—as in higher eukaryotes—require the interplay between the actin andMT cytoskeletons and landmark proteins atthe cortex (Siegrist and Doe 2007; Li and Gun-dersen 2008). This rather complex arrangementof components may be necessary to guaranteerobust polar growth. Coordinated oscillationsof actin polymerization, exocytosis, and Ca2+
levels associated with cell growth seem to be aconserved phenomenon shared among variousorganisms, including fungi (Das et al. 2012),mammalian cells (Wollman and Meyer 2012),and plant root hairs (Monshausen et al. 2008)and pollen tubes (Kroeger and Geitmann 2012).The oscillatory cell growth allows cells torespond more quickly and often to internaland external cues such as chemical or mechan-ical environmental signals.
56 N. Takeshita and R. Fischer
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